Assuming 1:1 conversion of RNA to cDNA is a fairly common convention. It isn't necessarily true, of course, but in lieu of any meaningful way to test it, it suffices. Besides: use of appropriate reference genes renders the matter essentially moot.

25ng of cDNA seems like a lot, especially since you're only using 100ng of RNA. If you're doing your PCRs in triplicate, you'll only be able to measure expression of a single gene. You'll also be adding a lot of cDNA synthesis reaction buffer to your PCR, which in my experience can hinder the PCR reaction.

I usually use ~8ng per well, from a prep made with 800ng of RNA, diluted down to give ~4ng.ul-1.

This means I can do a good thirty or so genes (in triplicate) if need be, and for many genes I could use 4ng or less.

End of the day, you don't want to add so much RNA that your cDNA synthesis is inefficient, you want your RNA to be clean (260/230 ratio >1.7 at the very least) so that you don't impair your cDNA synthesis, but beyond that, the input cDNA doesn't matter too much (though keep it consistent between samples if possible).

In your situation, I would use more RNA in each cDNA synthesis reaction (easily 10x as much), then dilute it down to ~5ng.ul-1, and then try this in test reactions with 1, 2, 3 and 5ul of cDNA per well (for instance).

agree with @ John Hildyard

regards

Don't bother with the "RNA input optimization" for making your cDNA. What you DO need to worry about is knowing the amount of cDNA that you are using in each qPCR reaction (I'm assuming you are looking at absolute expression values and not relative). Careful measuring is critical to reduce the variation due to experimental error.

You do need to calculate the efficiency of your qPCR primers using a standard curve. Read through the MIQE guidelines and discuss with a colleague before you continue your experiment, but it sounds like you are on the right track.

I agree with Jon Hildyard that it sounds like you are adding way too much cDNA in each reaction, unless your gene of interest has a very, very low expression. Good luck!

Check this link may be helpful

https://www.thermofisher.com/us/en/home/brands/thermo-scientific/molecular-biology/molecular-biology-learning-center/molecular-biology-resource-library/basic-principles-rt-qpcr.html

Greeting

Thanks for the help everyone.

According to the Thermofischer workflow guide, the RNA input seems to be merely to validate that the RT step in this procedure is working as it should. Its basically a validation step. (See Validation2, pg 6-10). : https://www.thermofisher.com/content/dam/LifeTech/Documents/PDFs/cms_075428.pdf

I think I might actually include this procedure. It should only take up 8 wells (4 wells in duplicates per gene tested), and use up 4 RT reaction mixes ( and those 4 cDNA samples can be used for a whole set of gene validations). The RNA I use will be pooled from all samples (a tiny bit from each).

I'll run the each of the 4 different RNA input cDNA samples next to the standards. If I can show that the amount of RNA I put in for each of my samples fits on the linear slope of the RNA input Validation plot, then I can safely say that the RT reaction is working as it should.

The only part of this that makes me feel that this RT reaction validation is not necessary is that I'm using such a small RNA input in the first place. If I was converting 500ng to 2000ng of RNA into cDNA, then I think this RT validation curve would be necessary. But using 100ng RNA input, I have doubts that the RT reactiion would ever be affected.